Extracellular Matrix Glycation and Crosslinking in Mammary Tumor Progression

Abstract

Breast cancer progression is accompanied by profound extracellular matrix (ECM) remodeling. A greater abundance of aligned fibrillar collagen is characteristic of invasive and aggressive breast cancers and has been associated with elevated activity of collagen crosslinking enzymes, such as lysyl oxidase (LOX) and lysyl hydroxylases (LH) and the formation of more mature collagen matrix crosslinks. Aligned collagen fibers can facilitate metastatic dissemination of tumor cells, and LOX inhibitors have been used to inhibit tumor progression and metastasis in experimental models. Thus, a better understanding of how matrix crosslinking alters tumor cell phenotypes, and behaviors would improve our ability to effectively treat aggressive metastatic breast cancer. Herein described is an experimental approach to glycate and crosslink a collagen-I/basement membrane extract ECM to study the impact of ECM crosslinking on mammary tumor progression in vivo. Moreover, glycation of collagen by sugars to form advanced glycation end products naturally occurs during aging, extending the potential relevance of this approach to research on mechanisms of aging involved in disease progression.

1. Introduction

Mammary tumor progression is accompanied by extensive ECM remodeling [1]. The most aggressive hormone receptor negative molecular subtypes of breast cancer present an abundant, aligned, and stiffened collagen matrix when compared to less aggressive hormone receptor positive manifestations of the disease [2, 3]. A stiff ECM can arise due to enhanced enzymatic crosslinking of collagen through lysyl oxidase (LOX), lysyl oxidase–like (LOXL) enzymes, and lysyl hydroxylases, and these modifications are implicated in tumor cell invasion and dissemination [4–8]. Indeed, stromal reservoirs of LOX and LOXL enzymes from fibroblasts and tumor-infiltrating macrophages can promote this metastatic process [3, 9, 10]. Similarly, glycation of collagen to form advanced glycation end products (AGEs) generates crosslinks in a naturally occurring phenomenon associated with aging and diabetes [11, 12]. AGEs have several known receptors whose stimulation can promote malignant progression and metastasis [13, 14]. However, the mechanisms by which ECM crosslinking and stromal stiffness alter the phenotypes of different tumor and stromal cell populations to foster aggressive tumor behaviors are still being elucidated.

Patient-derived xenograft tumor tissues have been exceptionally advantageous for studying tumor progression and clinically relevant responses to therapy as they demonstrate high fidelity to the genetic alterations and behavioral phenotypes observed in the originating patient tumor [15, 16]. This protocol describes a technique to glycate and crosslink collagen-I to form a stiffened collagen-I/basement membrane extract (BME) hydrogel ECM environment for breast-cancer-patient-derived xenograft (PDX) implantation. Prior work measuring the mechanical properties of collagen-I with varying levels of glycation and collagen concentration by rheometry informed this method [5, 17, 18]. However, good practice would necessitate testing and verification of each formulation by rheometry before performing orthotopic animal procedures. The approach entails preparation of the ribose crosslinked collagen I/BME hydrogels, the mouse surgical procedure area and materials, and the patient-derived xenograft fragments, prior to orthotopic injection of ECM hydrogels and implantation of the PDX fragments. This method can be applied to investigate the effects of matrix crosslinking and stiffening on overall tumor growth and metastasis, as well as to study the altered phenotypes and behaviors of unique stromal and tumor cell populations at different stages of tumor progression or in response to standard or novel treatment strategies (Fig. 1).

Fig. 1. Orthotopic implantation of breast-cancer-patient-derived xenograft tissue fragments with glycated collagen-I/BME hydrogels.

An inverted Y incision is made in the skin of the abdominal area. A skin flap is pulled back on one side, and the connection between the #4 and #5 mammary glands is severed with a cauterizing pen. The inguinal lymph node (#4 mammary gland) is located, and the mammary gland from the lymph node to the nipple (including the lymph node) is removed. A 50 μL volume of collagen-I/BME hydrogel (Col1/BME) is injected into the remaining portion of the mammary fat pad. If both #4 mammary glands will be implanted, this process is repeated on the opposite mouse flank. Returning to the first cleared and Col1/BME injected fat pad after waiting at least 3 min for the hydrogel to warm at the site of injection, thin tip cross-action forceps are used to make a pocket in the clear mammary fat pad. A patient-derived xenograft (PDX) tissue fragment is then placed over the mammary fat pad, and forceps are used to push the fragment into the premade pocket. Numbers (#) 1–5 indicate the positions of the mouse mammary glands. LN = inguinal lymph node

2. Materials

2.1. Preparation of Ribose-Crosslinked Collagen-I/BME Hydrogels

1. High-concentration rat tail collagen-I (concentration varies, typically 8–10 mg/mL).

2. 1 N sodium hydroxide (NaOH).

3. 2500 mM solution of L or D-ribose: For a 1 mL solution, weigh 0.188 g of ribose powder, and reconstitute in 0.1% glacial acetic acid in purified water.

4. pH strips.

5. 1.5 mL Eppendorf or 15 mL falcon tubes.

6. 10x DMEM/F12 media.

7. Basement membrane extract (BME, i.e., Matrigel).

8. 1 mg/mL fibronectin stock solution.

9. Micropipettes and tips.

2.2. Mouse Surgical Procedure Preparation

1. Electric razor appropriate for shaving mice.

2. Analgesics according to institutional protocol: Typical analgesics for a mouse weighing 25 g might include buprenorphine (50 μL of 25 μg/mL solution by intraperitoneal injection), meloxicam (100 μL of 1.25 mg/mL solution by intraperitoneal injection), and lidocaine (~30 μL 0.5% solution applied topically to surgical site).

3. Isofluorane for rodent anesthesia.

4. Isoflurane machine kit: isoflurane vaporizer, charcoal absorbent cannisters, induction chamber, nose cone adaptor, oxygen tank with regulator, and associated tubing.

5. Surgical tools: blunt tip scissors, pointed tip scissors, tissue forceps (straight with teeth), dressing forceps (serrated, bent), dressing forceps (serrated, straight), and forceps (thin-tipped cross action).

6. Wound clips (7 mm), wound clip applicator, and wound clip remover.

7. Battery-operated cauterizing pen and fine cauterizing tips.

8. Glass bead sterilizer.

9. Autoclave pouches for instruments and autoclave access.

10. Sterile saline solution (phosphate buffered saline).

11. Styrofoam surgical surface (large enough for a mouse with limbs extended).

12. Cotton-tipped applicators.

13. 70% ethanol.

14. Betadine scrub.

15. Clean mouse holding cages for post-surgery recovery monitoring.

16. Temperature-controlled heating pads (two or more) for animal surgery and recovery.

2.3. Patient-Derived Xenograft Preparation

1. Disposable scalpels (x2).

2. Surgical forceps and scissors (see Subheading 2.2).

3. Cryovials (optional).

4. Patient-derived xenograft tissues.

5. Sterile DMEM/F12 supplemented with 10% FBS.

6. Sixty-mm tissue culture dishes (as needed).

7. Carbon dioxide tank and chamber (for mouse sacrifice as needed).

2.4. Orthotopic Mammary Fat Pad Injection of ECM Hydrogels and Implantation of Patient-Derived Xenograft Tissues

1. Surgical tools and materials (see Subheading 2.2).

2. 27- or 26-gauge needles (5/8 inch)

3. 1 mL syringes

4. Ophthalmic ointment.

5. Laboratory tape.

6. Absorbent padding to cover surgical surface area.

7. Push pins.

8. Kitchen cling film for animal draping.

9. Mouse ear hole puncher or numbered metal ear tags and applicator (optional).

3. Methods

3.1. Preparation of Ribose-Crosslinked Collagen I/BME Hydrogels

Keep all reagents on ice unless otherwise specified.

1. Prepare stock 2500 mM solution of L or D ribose in 0.1% glacial acetic acid in purified water (see Note 1). Filter sterilize (0.2 μm pore size).

2. First, determine the amount of high concentration rat tail collagen-I (see Note 2) needed for double the number of injections planned (see Note 3). Add 2500 mM solution to the collagen-1 hydrogel to a final concentration of 500 mM (1:5 dilution). Note the initial stock concentration of collagen-I, and calculate the new collagen-I concentration following addition of the ribose solution. To prepare a non-glycated and crosslinked hydrogel control condition, generate a second lot of collagen-I with an equal volume of 0.1% acetic acid in purified water without ribose (1:5 dilution).

3. Incubate the collagen-I/0.1% glacial acetic acid +/− ribose solution for at least 14 days at 4°C to enable significant glycation and crosslinking prior to preparation of collagen-I/BME hydrogel mixtures (see Note 4).

4. Following the minimum incubation time for collagen-I solutions +/− ribose, prepare the collagen-I/BME hydrogels according to Table 1 (example calculations), except for the addition of the 1 N NaOH. The suggested formulation includes 20% BME (v/v), 10x DMEM/F12, and PBS up to the final volume (see Note 5).

5. Add half the calculated volume of the 1 N NaOH. Mix the collagen-I/BME solution by pipetting up and down slowly to prevent the introduction of air bubbles. Ensure that the hydrogels with (crosslinked) and without ribose (non-crosslinked control) are the same color (phenol red pH indicator), and verify that the mixtures have a similar pH using a pH strip. Add more 1 N NaOH as needed a few microliters at a time followed by mixing to achieve the desired pH (see Note 6).

6. Keep collagen-I/BME hydrogels on ice until mice are ready for orthotopic injection.

Table 1.

Example calculations for the preparation of glycated collagen-I/BME hydrogels

Reagents
Stock collagen concentration (mg/mL)	10
Desired collagen concentration (mg/mL)	4.8
Stock Ribose concentration (mM)	2500
Desired Ribose concentration (mM)	500
Desired BME (Matrigel) concentration (%)	20
Experimental setup	Soft	Stiff
Injection volume (μL)	50	50
Number of injections (#glands x 2)	20	20
Total volume	1000	1000
Step 1: Collagen glycation	Soft	Stiff
Component	Volume (μL)	Volume (μL)
Collagen I (rat tail)	800	800	(Collagen I)
0.1% acetic acid	200	–	8.00
0.1% acetic acid +2.5 M ribose	–	200	Mg/mL
Total volume	1000	1000
Incubate @ 4 °C for 14 days
Step 2: Collagen preparation	Soft	Stiff
Component	Volume (μL)	Volume (μL)	(Collagen I)
Collagen I ± ribose	600	600	4.8
BME (Matrigel)	200	200	Mg/mL
1 N NaOH	14	14	(BME)
10X DMEM	100	100	20.00
1X PBS	86	86	%
Total volume	1000	1000

3.2. Preparing for Mouse Surgical Procedures

Mice should be 3 weeks of age to ensure that the endogenous mammary epithelium can be efficiently cleared from the inguinal mammary fat pad. Ensure that the preparation and use of anesthesia equipment and surgical areas adhere to specific approved Institutional Animal Care and Use Committee protocols.

1. Autoclave surgical instruments and wound clips at least 1 day prior to performing mouse surgery to ensure sterile materials are ready before use.

2. Shave the abdominal area where the skin incisions will be made in a designated area that is separate from where the surgical procedure will be performed.

3. Optional: Intraperitoneal injection of analgesics can be performed at this time, prior to induction of isofluorane anesthesia. Follow an approved animal research protocol. Suggested analgesics (and their concentrations) are provided.

4. Set up isofluorane anesthesia machine. Ensure all tubing is connected securely: Isofluorane vaporizer connected to the dual diverter valve that connects to the induction chamber and the nose cone adaptor. Check that exhaust tubing from both the induction chamber and the nose cone adaptor is directed to charcoal filtration canisters. Weigh canisters before use, and replace if they exceed the predetermined and recorded maximum weight. Keep valves to the chamber and nose cone supply closed if not in use to protect mice and limit room air pollution. Verify that there is sufficient isofluorane in the vaporizer by looking at the level through the glass window on the front of the machine. Top up isofluorane to the fill level as needed. Briefly open flow of oxygen tank to ensure all connections are tight and that the tank regulator displays sufficient oxygen pressure to perform the required surgeries. Replace the oxygen tank if the pressure gauge reads less than 500 PSI.

5. Set up a surgical area. Sanitize the surgical surface of a bio-safety cabinet with chlorhexidine solution or Clidox. Place temperature-controlled heating pads beneath half of a clean recovery cage nearby, where mice will be monitored post-surgery. Leaving half the cage off the heat source will allow mice to escape the heat if needed. Tape a thin Styrofoam platform with absorbent material covering the surface that is large enough to accommodate a mouse onto the heating pad in the surgical area. This creates a thin barrier between the heat source and the animal. Affix the nose cone to this Styrofoam surgical table and drape the surface with absorbent padding.

6. Prepare a sterile surface for using the sterile tip technique for all surgical tools. If an autoclave pouch was used for sterilizing instruments, then the inner surface of the pouch can serve this purpose. Place the tips of surgical tools (scissors, two forceps, thin tip cross-action forceps, would clip applicator, etc.) on the sterile surface.

7. Place remaining surgical materials near the surgical area (cauterizing pen, ophthalmic ointment, cotton-tipped applicators, 70% ethanol, betadine scrub, sterile saline solution).

8. Plug in and turn on a glass bead sterilizer for cleaning surgical tools in-between animals. Place the bead sterilizer in a convenient location near the planned surgical procedure area.

3.3. Patient-Derived Xenograft Preparation

1. If transplanting fresh breast cancer PDX tissue from a live animal, first sacrifice the animal using an approved humane protocol, such as carbon dioxide inhalation followed by cervical dislocation.

2. Excise the tumor, and place the tumor into a 60 mm tissue culture dish containing 3–4 mL of DMEM/F12 media supplemented with 10% FBS. Chop the tumor into large fragments (5 × 5 mm). At this point, some of the tumor may be cryopreserved by placing fragments into a cryovial with 95% FBS and 5% DMSO and slowly freezing the vial in a freezing container on dry ice before subsequent transfer to −80°C.

3. If transplanting viably cryopreserved PDX tissue as above, place the tumor fragment contents of a thawed cryovial into a 60 mm tissue culture dish containing 3–4 mL of DMEM/F12 media supplemented with 10% FBS.

4. Using scalpels and forceps as required to secure the tissue, chop PDX tumor fragments into smaller 2 × 2 mm pieces in the 60 mm dish. These fragments are now ready for transplantation (see Note 7).

3.4. Orthotopic Mammary Fat Pad Injection of ECM Hydrogels and Implantation of Patient-Derived Xenograft Tissues

Ensure that all animal-related procedures adhere to specific approved Institutional Animal Care and Use Committee protocols. Animal surgeons should wear appropriate personal protective equipment, use sterile tip technique for surgical instruments throughout the procedure and a glass bead sterilizer to re-sterilize instruments between animals. Ensure instruments are sufficiently cooled before contacting animals.

1. Load 1 mL syringes with remaining analgesics and prepared collagen/BME mixtures with and without ribose-mediated glycation and crosslinking (see Note 8). Use 27- or 26-gauge needles (5/8 inch) at the end of the syringes (see Note 9).

2. Turn on oxygen flow and set oxygen flow regulator to ~1.5 L/min.

3. Place first mouse in the induction chamber and set isofluorane vaporizer to ~3%. Monitor mouse until fully anesthetized with a regular, reduced breathing rate.

4. Open the valve controlling flow of anesthesia to the nose cone. Transfer mouse to the nose cone from the induction chamber and close the valve directing flow to the chamber.

5. While the mouse is on its stomach, lubricate eyes with ophthalmic ointment using a cotton-tipped applicator to prevent eyes from drying out during the procedure.

6. If needed, apply an ear tag or use an ear puncher to number mouse accordingly. Then flip the mouse onto its back while maintaining its nose in the nose cone adaptor. Use a toe pad pinch to check for a reflex reaction in the mouse. Monitor animal’s vital signs, such as respiratory pattern, skin/mucous membrane color, and depth of anesthesia. Adjust the flow of anesthesia until mouse insensitivity to pain is confirmed.

7. Apply laboratory tape over the feet to secure the mouse to the surgical area surface with forelimbs and hindlimbs pointing up and down, respectively, away from the body at about 45° angles.

8. Drape the surgical area with a sterile material. Typical kitchen cling film provides an appropriate sterile material for this purpose. Cut an opening in the draped cling film away from the surgical area, and clean the surface of the skin using cotton-tipped applicators with three alternating swabs each of betadine scrub and 70% ethanol. Draping will help prevent contamination of the surgical site from non-sterile parts of the animal.

9. In the center of the abdomen, between the nipples of the #4 mammary fat pad, pick up the skin with forceps, and snip a tiny hole in the skin with surgical pointed tip scissors large enough to insert the tip of the blunt tip scissors. Slide the blunt tip scissors into the opening, and push the end of the scissors under the skin toward the head of the mouse to separate the skin from the peritoneal lining. Using the same scissors, make an initial vertical midline incision starting from the small initial opening in the skin toward the head of the mouse about 1 cm in length (see Note 10).

10. Then, in the same way, use the surgical scissors to make two lateral incisions in the skin about 1 cm or less in length in the direction of each hindlimb and between the nipples of the #4 and #5 mammary fat pads. The three incisions should now form an inverted Y. At this point, an appropriate volume of topical analgesic can be added dropwise from a syringe to the site of the skin incisions.

11. Peel back a triangular flap of skin on one flank of the animal, pulling the skin with forceps and pushing the abdomen away in the opposite direction with a cotton-tipped applicator dipped in sterile saline solution as needed to fully uncover the #4 mammary fat pad.

12. Use a push pin to pin the flap of skin down to the Styrofoam surgical surface. Angle the push pin away so that it holds the skin flap in place and does not obstruct access to the skin opening.

13. Locate the connection between the #4 and #5 mammary fat pads near the point where the hind limb meets the abdomen, pull up on this section of the mammary gland with forceps, and sever this connection using a cauterizing pen (see Note 11).

14. Locate the major blood vessels and inguinal lymph node located approximately in the middle of the #4 mammary fat pad. Grab the #4 mammary fat pad toward the outside of the mammary gland from the lymph node, and use the surgical scissors to cut the mammary gland on the abdominal side of the lymph node completely through the fat pad without cutting the skin underneath. Then gently pull the mammary gland held by the forceps away from the body, and use the scissors to cut it away from the inner surface of the skin until the entire mammary gland from the lymph node to the nipple has been excised (see Note 12). The #4 mammary fat pad is now cleared of its endogenous epithelium.

15. Take a syringe loaded with collagen-I/BME mixture in one hand, and with the other hand, hold and pull gently on the remaining portion of the #4 mammary gland. Stretching the gland slightly will aid with initial insertion of the needle for the loaded syringe. Ensure that the bevel of the needle is entirely submerged in the fat pad, and lift up gently on the syringe to see the needle tip, and make sure that it has not been inserted through the back of the mammary fat pad. Then relax the mammary gland tension with the forceps, and begin injecting the collagen-I/BME hydrogel until a total volume of 50 μL is injected (see Note 13). A small bubble should be visible in the remaining portion of the mammary gland. Remove the push pin holding the skin down, and gently replace the skin flap over the excised and injected mammary gland.

16. Repeat steps 13–15 for the opposite flank (#4 mammary fat pad).

17. Return to the first fat pad that was cleared and injected with collagen I/BME hydrogel. Re-pin the skin flap down to the Styrofoam surgical area surface as before with a push pin. Take the thin tip cross-action forceps in one hand, and with the other hand, use additional forceps to pick up the remaining #4 mammary gland, and pull gently to apply some tension as described above. Pierce the mammary gland with the forceps, then relax gland tension, and squeeze the cross-action forceps slightly to open a small pocket in the mammary gland.

18. With forceps, take a small fragment of PDX tumor material (2 × 2 mm) that is resting in a medium bath of DMEM/F12 supplemented with 10% FBS (see Subheading 3.3).

19. While still holding the mammary gland with forceps in one hand, place the PDX fragment on the mammary gland at the site where the pocket was made in step 17. Then use the forceps to gently nudge the PDX fragment into the pocket (see Note 14). The fragment will be engulfed in the pocket and most likely be visible within the hydrogel due to the contrast in appearance. Remove the push pin, and replace the skin flap gently over the implanted mammary gland.

20. Repeat steps 18–19 for the cleared and hydrogel-injected mammary gland on the opposite flank of the mouse.

21. Apply wound clips to close the inverted Y skin incisions. Using forceps in each hand, pick up the skin at the bottom of the vertical incision, and reposition the skin so that it can be held together in one pair of forceps. With your newly freed hand, use the would clip applicator to apply a single wound clip closing the vertical incision. Apply multiple would clips as necessary, being sure to leave a small space between wound clips and that the clips are not so tight that they risk strangulating the skin or impairing mouse mobility. Repeat this closure process for the two lateral skin incisions of the inverted Y shape.

22. Place the mouse in the recovery cage on a heating source in a place where it can easily be monitored for recovery to full mobility.

23. Return mice to their home cage once fully recovered from the procedure. Administer additional analgesics post-surgery according to approved practices stipulated in your specific research animal use and care protocol.

24. Repeat this procedure for the desired number of mice with equal numbers of injections/tumor implantations performed for collagen I/BME hydrogels with and without ribose-mediated crosslinking for effective comparison (see Note 15).

4. Notes

1. The use of L or D ribose have biological implications. Naturally occurring D ribose may have more biological activity and be more relevant to physiological collagen processing and aging mechanisms. Alternatively, L ribose may form more biologically inert glycation products such that phenotypes can be attributed to the effects of enhanced crosslinking rather than collagen glycation.

2. High-concentration rat tail collagen stock concentrations vary, but most preparations fall between 8 and 10 mg/mL concentration. Another important consideration is the potential immunogenic impact of rat tail collagen. An alternative would be to use mouse tail collagen either prepared in the laboratory or purchased commercially.

3. Due to the viscosity of these collagen/BME mixtures, it is recommended to prepare double the amount needed because of loss of volume with pipette and syringe/needle manipulations.

4. The collagen should be maximally glycated and crosslinked after 14 days. Note that these collagen preparations are stable for at least 3 months at 4°C so that they can be incubated well in advance of planned mouse orthotopic injections.

5. The specific formulation of these collagen I/BME hydrogels is suggested as a tested starting point. It would also be conceivable to test and study the impact of different collagen-I and BME concentrations as well as the addition of different ECM components within this system.

6. Experience with these hydrogels has shown that it is better to have a pH slightly toward the basic (orange color) side of the spectrum as more acidic pHs (pink color) are deleterious to cell viability and growth. The pH of collagen I/BME hydrogel mixtures should be tested using pH strips before proceeding with animal injections.

7. The dish of prepared PDX fragments should be kept on ice if surgeries will continue for an extended period. The size of tumor tissue fragments suggested is a recommendation for ease and success of tumor implantation. Variations could be performed with larger fragments or multiple fragments as desired, with the stipulation that equal tumor volumes are implanted for comparison of experimental conditions.

8. It is suggested to load only enough volume for 4–5 injections of collagen I/BME hydrogels at a time to avoid the hydrogels sitting in the syringe for long periods of time. However, allowing the hydrogel in the syringe to warm to room temperature or warming it with your fingers or a heating pad prior to injection will help ensure that the hydrogel is retained in the fat pad with minimal leaking.

9. Total 26- or 27-gauge needles are optimal for this procedure. It is not recommended to use needles with a lower or larger gauge, because the hydrogels may solidify and occlude needles with a lower gauge as they warm, while needles with a larger gauge may induce more leaking upon hydrogel injection.

10. Keeping these incisions as small as possible will enable cleaner wound closure with improved comfort and recovery for the mice. However, incisions can be made larger to ensure successful hydrogel injection and tumor implantation. Blunt-ended scissors are optimal for this step as they reduce the risk of piercing the abdomen, skin, or other mouse tissues and organs. Also note that this procedure accounts for injecting/implanting both #4 mammary glands of one animal. If only one #4 mammary gland will be injected/implanted, then only one vertical incision at a 45° angle in the direction of the #4 nipple on the intended side is required to follow the remaining steps.

11. If scissors are used in place of a cauterizing pen to sever the connection between the #4 and #5 mammary fat pads, extensive bleeding may occur which will lower the likelihood of mouse survival following surgery. Therefore, the use of a cauterizing pen for this process is strongly recommended.

12. Following removal of the nipple proximal portion of the mammary gland, the researcher should visually check that no remaining fragment of mammary gland is attached to the inner surface of the skin in the region. Moreover, these sections of cleared mammary fat pad can be saved and mounted onto a slide for staining with hematoxylin or carmine to visualize the ductal tree of the mammary epithelium with its terminal end buds. In this way, successful clearing of the endogenous epithelium can be verified.

13. Following injection of hydrogels, a waiting period of at least 3 min is suggested to allow the hydrogel time to warm and begin to solidify prior to implantation of PDX fragments. Allowance for this additional time has enhanced outcomes for the following steps in this method.

14. It is most often more effective to use closed forceps to push the PDX fragment into the premade mammary fat pad pocket, as opposed to holding the fragment and placing it in the pocket. Extra care should be taken to avoid displacing the injected hydrogel in this step. Practice may be helpful here to improve efficiency and frequency of success.

15. This procedure could also be adapted in a number of ways. While thus far unsuccessfully employed for PDX mammary tumors, an alternative approach would be to resuspend and disperse dissociated tumor cells or tumor cell lines within the collagen I/BME hydrogels prior to orthotopic injections. This procedure could also be adapted for injection/implantation of additional types of tumor cells/PDX tissues at other locations in the mouse, such as the pancreas, prostate gland, subrenal capsule, or subcutaneous sites.